Name: visualizing stromule frequency with fluorescence microscopy
Uploaded: 2016-11-23T14:00:00.000+00:00
Duration: 8 min 27 s
Description: Watch this Scientific Journal Video about Visualizing Stromule Frequency with Fluorescence Microscopy at JoVE.com

J.O.B. and A.M.R. were supported by predoctoral fellowships from the National Science Foundation.

NOTE: For this protocol, we have focused on assaying stromule frequency in the epidermis of N. benthamiana leaves. Several stable transgenic lines have been generated that can be used for this purpose, including 35SPRO:FNRtp:EGFP13 and NRIP1:Cerulean6. Both of these lines show robust expression of fluorophores in the chloroplast stroma of leaves grown under a wide range of conditions. Alternatively, chloroplast-targeted fluorophores may be transiently expressed in N. benthamiana using Agrobacterium transformations13. This is less ideal than the transgenic lines, since Agrobacterium infiltrations induce some basal defense responses in N. benthamiana and interactions with Agrobacterium can alter stromule frequency in the leaf14, potentially complicating interpretation of results. Finally, to visualize stromule formation in vitro, chloroplasts may be extracted from any plant species, using either genetically encoded fluorophores or a fluorescent dye, as described in section 5 below.9,15
NOTE: Detailed methods for plant cultivation have been previously described.16 Briefly, grow N. benthamiana plants in 4&#34; pots filled with any professional soil mix that provides good drainage. Cover seedlings with a clear plastic dome for the first 10-14 days to provide a humid environment for germination. Add any standard fertilizer mix following manufacturer&#39;s instructions to 14-day-old plants. Grow plants under white light, using ~100 &#181;mol photons m-2 sec-1 light intensity. Water plants regularly.
1. Preparing Leaf Samples for Visualization
NOTE: Stromule dynamics are affected by wounding8, so tissue preparation should be conducted immediately before visualizing stromules by confocal fluorescence microscopy. Ideally, a sample should be visualized with 15 min after removal from the plant.
<ol>
	<li>Cut a small section of the leaf using a very sharp razor blade. Always use the same region of the leaf for consistency, and be certain to visualize the same surface (adaxial or abaxial) in all experiments.</li>
	<li>Immediately after cutting the leaf section, submerge the leaf section in a 5 ml needleless syringe filled with water, remove air from the syringe, and apply a vacuum by covering the syringe orifice with a finger and pulling the plunger.
	<ol>
		<li>Release the plunger gently to avoid damaging the leaf section. Repeat this two or three times, or until most of the air has been removed and the leaf appears to be deep green. 
		NOTE: This step removes air in the leaf that can interfere with accurate visualization of stromules.</li>
	</ol>
	</li>
	<li>Add a drop of water to a slide and place the leaf section on this drop. Then, add another drop of water to the top of the leaf, and place a cover slip over the leaf. If there are any air bubbles, gently tap the cover slip until they are removed. The slide is now ready for microscopy, and should be visualized immediately.</li>
</ol>
2. Visualizing Stromules with Confocal Fluorescence Microscopy
<ol>
	<li>With transmitted light, focus on a field of view near the center of the leaf section (away from the cells damaged by the razor blade). Use a 20X objective so that many chloroplasts can be visualized simultaneously. Save an image with transmitted light so that cell types can be distinguished later, if necessary.</li>
	<li>Switch the illumination source to a laser with appropriate excitation and emission filters for the fluorophore being used. For example, excite GFP with a 488 nm laser and collect emissions between 500 nm and 525 nm.
	<ol>
		<li>Using the microscope&#39;s software, select the GFP channel and click to adjust the pinhole aperture to 1 Airy unit. Select laser scanning, to start visualizing the sample, and then click the GFP channel to adjust the laser intensity and the detector gain to minimize the laser power while still clearly visualizing stromules. Once these settings have been determined, keep them as consistent as possible throughout all experiments.</li>
	</ol>
	</li>
	<li>Prepare a z-stack experiment that will collect a series of images through the leaf epidermis in the field of view.
	<ol>
		<li>For the z-stack, include all epidermal chloroplasts in the field of view to avoid bias (for example, if chloroplasts near the leaf surface have more stromules under conditions tested).</li>
		<li>Take images at the maximum interval possible that will include all stromules but minimize the time required for a z-stack. For example, if 1 Airy unit is equivalent to an in-focus confocal section that is 2 &#181;m deep, taking an image every 2 &#181;m is likely ideal. 
		NOTE: The leaf epidermis is an uneven surface, so the total depth of the z-stack will vary depending on the sample; most z-stacks will require 10-20 images, but may include more.</li>
		<li>Adjust scanning speed and image resolution as necessary to make sure that the z-stack is collected quickly (typically within 5-10 min, if possible). Save the z-stack for later analysis.</li>
	</ol>
	</li>
</ol>
3. Image Processing
<ol>
	<li>Determine stromule frequency using any image analysis software. For this protocol, we recommend using ImageJ, which is publicly available from the National Institutes of Health (http://imagej.nih.gov/ij/), or the popular upgrade of ImageJ, Fiji (http://fiji.sc/Fiji).</li>
	<li>Merge the z-stack into a single image using the maximum intensity projection in the software.</li>
	<li>Identify and manually count all chloroplasts in the image.</li>
	<li>For each chloroplast, visually determine whether one or more stromules are seen extending from the chloroplast in the merged z-stack image. For examples, see Figure 1. 
	NOTE: Since the biological functions of stromules are not known, any thin, stroma-filled, tubular extension from the chloroplast can be considered a &#34;stromule&#34;, regardless of length. As a general rule, a stroma-filled extension from the chloroplast is a stromule if it is less than approximately 1 &#181;m in diameter along most of its length7. Ensure that one person analyzes all images to control for subjective differences in determining whether or not a chloroplast has a stromule. A second researcher should independently verify the stromule frequencies to further reduce subjective biases.</li>
</ol>
4. Experimental Design and Sampling
NOTE: Stromule frequency is highly variable between leaves, but several reports suggest that there is little variation in stromule frequency within an individual leaf9,17.
<ol>
	<li>Calculate the leaf stromule frequency from a single field of view of one leaf. Discard the leaf, and consider this an independent sample. Do not consider separate fields of view from one leaf as independent samples. 
	NOTE: There is no strong consensus on how to best measure changes in stromule activity, and until the function of stromules is more clearly defined, researchers should continue to be creative in quantifying stromule dynamics. For comparison across the literature, however, common standards of reporting should be used in addition to more creative approaches. At a minimum, always report the frequency of chloroplasts that have one or more stromules.</li>
	<li>Determine the frequency of chloroplasts that have stromules in a field of view. Use ImageJ to identify all chloroplasts in a field of view. Then, for each chloroplast, determine whether the chloroplast has formed at least one stromule. Report stromule frequency as the percentage of chloroplasts with a stromule. 
	NOTE: Some researchers transform the percentage, for example with the arcsine transformation, before reporting stromule frequencies; while this is an option for statistical analysis, the arcsine of stromule frequency is not an intuitive value, and does not need to be reported in the main text or figures of a study.
	<ol>
		<li>As an alternative approach, count the total number of stromules, and divide it by the total number of chloroplasts. 
		NOTE: Under most circumstances, this will yield similar results to approach 4.2; it may overestimate the stromule frequency, however, if a single chloroplast has formed many stromules. In the example shown under representative results, for example, several chloroplasts have two or more stromules, raising the number of stromules per chloroplast to 40/87 (versus a stromule frequency of 33/87).</li>
		<li>Alternatively, determine the stromule length instead of stromule frequency. Length can be measured in ImageJ by tracing the stromule with the freehand line tool. 
		NOTE: Since the biological role of stromules remains unknown, it is not clear that stromule length affects their function. Report significant differences in stromule length if they are observed, but also report the stromule frequencies (as described in 4.2). 
		NOTE: Stromule frequency can be highly variable among different leaves under the same growth conditions. Therefore, although determining stromule frequency can be time-consuming, experiments should be carefully designed to include large sample sizes. Using datasets from our lab, we determined that a sample size of at least 16 plants for each treatment is usually sufficiently powerful to determine if there is a statistically significant difference of at least a 1.5-fold change in stromule frequency between two conditions (with standard &#945; = 0.05 and &#946; = 0.80).</li>
	</ol>
	</li>
	<li>Statistical analysis of results.
	<ol>
		<li>To compare the mean stromule frequencies of two treatments, use the Welch&#39;s t test (also known as the heteroscedastic t test or t test assuming unequal variance). 
		NOTE: This test is not as powerful as the conventional Student&#39;s t test, but Welch&#39;s t test is advantageous because it does not assume that the variance in stromule frequency distribution is similar between treatments. 
		NOTE: Since stromule frequency is a &#34;proportion&#34;, its sampling distribution is predicted to be binomial rather than normal, which could theoretically skew statistical analysis if sample sizes are very low or if stromule frequency is very low (close to 0%) or very high (close to 100%). Some reports have used arcsine transformations before statistical analysis, which is intended to transform a binomial distribution to a normal distribution, and thus satisfy the standard requirements of a Student&#39;s t test or ANOVA. In practice, however, arcsine transformations have little or no effect on statistical analysis, and are therefore not recommended. Instead, repeat experiments to increase sample size, which will increase statistical power and reduce errors in interpretation of data.</li>
		<li>Whatever statistical approach is used, be sure to carefully report the sampling strategy, sample size, statistical test, and p value for each experiment. 
		NOTE: As with other fields of biology, a p value less than 0.05 may be considered &#34;statistically significant&#34;, although researchers should certainly report the precise p value obtained. If p is slightly above 0.05, increasing the sample size with two or three experiments may help to clarify whether or not the difference observed is reproducible and statistically significant.</li>
	</ol>
	</li>
</ol>
5. Extracting Intact Chloroplasts to Visualize Stromule Dynamics
NOTE: Several methods have been used to isolate chloroplasts from leaves, including a slightly different protocol in a recent study on stromule formation in vitro15. The protocol detailed below uses a relatively simple method that does not yield biochemically pure chloroplast samples, but does instead isolate a large quantity of intact, healthy chloroplasts9,18.
<ol>
	<li>Prepare cold extraction buffer: 50 mM Hepes NaOH, 330 mM sorbitol, 2 mM EDTA, 1.0 mM MgCl2, and 1.0 mM MnCl2. Adjust the pH to 6.9 with NaOH and HCl, and refrigerate before use. 
	NOTE: Although not necessary, using cold buffers and keeping samples on ice when possible will yield a higher proportion of intact chloroplasts.</li>
	<li>Prepare isolation buffer: 50 mM Hepes NaOH, 330 mM sorbitol, 2 mM EDTA, 1.0 mM MnCl2, 1.0 mM MgCl2, 10 mM KCl, and 1.0 mM NaCl. Adjust the pH to 7.6 with NaOH and HCl and refrigerate before use.</li>
	<li>If the leaves are not expressing a genetically encoded stromal fluorophore, prepare carboxyfluorescein diacetate (CFDA) solution: 50 mM CFDA stock solution in dimethyl sulfoxide (DMSO) (2.3 mg CFDA per 1.0 ml DMSO). 
	NOTE: The exact concentration is not critical. Keep CFDA stock in the dark at all times. Store small aliquots of 50 mM CFDA at -20 &#176;C.</li>
	<li>To isolate chloroplasts, remove ~5-10 g leaves from several plants and briefly rinse in cold water. Immediately transfer to 50 ml cold extraction buffer. Grind leaves using a blender with several short pulses. Filter through two or three layers of cheesecloth to remove leaf debris, divide the extracted chloroplasts into two 50 ml centrifuge tubes, and centrifuge for 1 min at 750 x g.</li>
	<li>Discard the supernatant and resuspend the green chloroplasts in 10 ml isolation buffer. Centrifuge again for 1 min at 750 x g, discard supernatant, and resuspend chloroplasts in isolation buffer to bring final volume to 5 ml.</li>
	<li>Transfer 20 &#181;l of the chloroplasts to a slide, cover with a coverslip, and begin microscopy. 
	NOTE: Chloroplasts from transgenic plants expressing plastid-targeted fluorescent proteins are now ready for visualization by confocal fluorescence microscopy.</li>
	<li>Stain chloroplasts from plants that are not expressing plastid-targeted fluorescent proteins to visualize stromules.
	<ol>
		<li>Add 5 &#181;l of 50 mM CFDA stock to the 5 ml chloroplasts-suspended in isolation buffer. Bring the final concentration to 50 &#181;M.</li>
		<li>Allow chloroplasts to incubate for 5 min, and then transfer a small aliquot (~50 &#181;l) to a slide, cover with a coverslip, and begin microscopy.</li>
		<li>Use a FITC or GFP filter set. Use a 20X objective to visualize many chloroplasts simultaneously, or a higher objective to visualize a single isolated chloroplast. 
		NOTE: CFDA will fluoresce inside the stroma of intact chloroplasts.</li>
		<li>If there is strong background fluorescence, wash the chloroplasts again in 10 ml isolation buffer after the 5 min incubation with CFDA, centrifuge for 1 min at 750 x g, discard supernatant, and resuspend in 5 ml isolation buffer.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

When investigating stromules, three important factors must be considered throughout: (i) manipulation of the plant tissue must be kept to an absolute minimum, (ii) the experimental system must be kept consistent, and (iii) sampling strategies must be carefully planned to ensure robust, reproducible data are analyzed.
Stromules are remarkably dynamic: they can extend and retract rapidly before an observer&#39;s eyes under the microscope. Moreover, stromule frequency varies significantly in response to a wide range of treatments, including stimuli that cannot be avoided in order to visualize stromules (such as leaf wounding). The primary solution to this problem, which is addressed with the protocols described here, is to conduct well-controlled experiments, keeping all variables consistent. This includes, for example, preparing each sample for visualization immediately before bringing it to the microscope; plants should not be damaged until immediately before visualization. The second solution to this problem is to minimize the complexity of protocols: ideally, any treatments should be applied directly to the plant without removing the leaf or causing any damage.
Stromules have been studied in a dizzying array of species and cell types, including wheat root hairs, tomato fruits, and etiolated tobacco seedlings8,19. These pioneering studies advanced the understanding of stromule dynamics during plant development and explored the range of stromule activity. Drawing comparisons from these various experimental systems, however, can lead to misinterpretations, since different types of plastids are involved in considerably different biological processes.
For example, chloroplasts make stromules in response to oxidative stress caused by the photosynthetic electron chain, but since leucoplasts lack the photosynthetic machinery, they are not sensitive to the same stimuli9. Any experimental system may be used to investigate stromules, but the experimental system must be kept consistent throughout the study if any comparisons are to be drawn. Factors to consider include species, cell type, developmental stage or age of the plant, and method of staining stromules (whether through stable transgene expression, transient transgene expression, or an external fluorophore); even slight variations of these factors can confound later interpretation of results.
Finally, any study of stromule biology must use a robust, reliable sampling strategy to ensure that results are reproducible and biologically relevant. Sampling repeatedly from a single plant, e.g., taking images from several regions of one leaf, will apparently improve statistical power, but can be misleading. As shown in the representative data above, the stromule frequency of a single leaf may vary dramatically from the mean stromule frequency across many leaves: in that leaf, which was visualized during the day, 37.9% of chloroplasts made stromules, although the mean stromule frequency across leaves during the day was almost half that, only 20.8%.
Moreover, given how little is known about stromule dynamics at this time, any experiment should be entirely replicated at least three times, and more if possible, to ensure that unforeseen variables are not responsible for any changes in stromule activity that are observed. Above all else, since the stromule biology field is just beginning to take off, researchers should carefully document and report all aspects of experimental design, including creative deviations from the simple protocol outlined here.

Chloroplasts are dynamic organelles in plant cells responsible for photosynthesis and a host of other metabolic processes. Signaling pathways from the chloroplast also exert significant influence on plant physiology and development, coordinating plant responses to environmental stress, pathogens, and even leaf shape1-6. Recently, biologists have gained interest in a poorly understood aspect of chloroplast structure: stromules, very thin stroma-filled tubules that extend from the surface of the chloroplast7.
The biological functions of stromules remain unknown, although stromule frequency is known to vary in response to environmental stimuli7-9, and stromules may be capable of transmitting signaling molecules between organelles6. All types of plastids (not only the green, photosynthetic chloroplasts, but also clear leucoplasts, starch-filled amyloplasts, and pigmented chromoplasts, to name a few types of plastids) make stromules, and stromules are found in all land plant species that have been examined to date. Stromules can extend and retract dynamically, appearing or disappearing within seconds, or they can remain relatively stationary for long times. One of the major hurdles facing stromule biologists is that stromules are often studied using dramatically different methods, tissues, and species, making comparisons across the stromule biology literature difficult. Going forward, standard practices and thorough descriptions of the experimental systems used to study stromules will be critical to discovering the function of these ubiquitous features of chloroplast morphology.
Here we describe methods for visualizing stromule formation in the epidermal chloroplasts of Nicotiana benthamiana leaves. In the mesophyll, chloroplasts are densely packed into large, three-dimensional cells, which makes it difficult to accurately and rapidly visualize stromules by confocal microscopy. By contrast, epidermal cells are relatively flat, contain fewer chloroplasts, and are at the surface of the leaf, allowing for easy and rapid visualization of stromules. N. benthamiana is an ideal model system for these experiments because, unlike many plant species, all cells in the epidermis of N. benthamiana make chloroplasts10. In the epidermis of most plants, including Arabidopsis thaliana, only the stomatal guard cells have chloroplasts, while other epidermal cells have "leucoplasts", plastids that are clear, relatively amorphous, and nonphotosynthetic9,11,12. Thus, whereas a single field of view of an A. thaliana epidermis might show only a handful of chloroplasts in a pair of guard cells, a field of view of an N. benthamiana epidermis will include dozens or even hundreds of chloroplasts. All of the methods described here, however, can be modified to investigate other questions in stromule biology; for example, we have used the same approach to study leucoplast stromules of A. thaliana9.

<table><tbody><tr><td>Hepes</td><td>Sigma-Aldrich</td><td>H3375</td><td></td></tr><tr><td>NaOH</td><td>Fischer-Scientific</td><td>S320-1</td><td></td></tr><tr><td>Sorbitol</td><td>Sigma-Aldrich</td><td>S1876</td><td></td></tr><tr><td>EDTA</td><td>Fischer-Biotech</td><td>BP121</td><td></td></tr><tr><td>MnCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>221279</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>M0250</td><td></td></tr><tr><td>KCl</td><td>Sigma-Aldrich</td><td>P3911</td><td></td></tr><tr><td>NaCl</td><td>Sigma-Aldrich</td><td>S9625</td><td></td></tr><tr><td>Laser Scanning Confocal Microscope&amp;nbsp;</td><td>Carl Zeiss Inc</td><td>Model: LSM710</td><td></td></tr><tr><td>Carboxyfluorescein diacetate (CFDA)</td><td>Sigma-Aldrich</td><td>21879&amp;nbsp;</td><td></td></tr><tr><td>Dimethyl sulfoxide (DMSO)</td><td>EMD</td><td>MX1458-6</td><td></td></tr><tr><td>Waring blender</td><td>Waring&amp;nbsp;</td><td>Model: 31BL92</td><td></td></tr><tr><td>Fiji</td><td>fiji.sc</td><td></td><td>Open-source software for analyzing biological images</td></tr></tbody></table>

visualizing stromule frequency with fluorescence microscopy

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but whose functions remain mysterious. Alongside growing attention on the role of chloroplasts in coordinating plant responses to stress, interest in stromules and their relationship to chloroplast signaling dynamics has increased in recent years, aided by advances in fluorescence microscopy and protein fluorophores that allow for rapid, accurate visualization of stromule dynamics. Here, we provide detailed protocols to assay stromule frequency in the epidermal chloroplasts of Nicotiana benthamiana, an excellent model system for investigating chloroplast stromule biology. We also provide methods for visualizing chloroplast stromules in vitro by extracting chloroplasts from leaves. Finally, we outline sampling strategies and statistical approaches to analyze differences in stromule frequencies in response to stimuli, such as environmental stress, chemical treatments, or gene silencing. Researchers can use these protocols as a starting point to develop new methods for innovative experiments to explore how and why chloroplasts make stromules.

This protocol was used to visualize stromule frequency at day and at night in the cotyledons of young N. benthamiana seedlings. Slices from a z-stack were merged into a single image (Figure 1A). For visual purposes, that image was then desaturated and inverted so that stroma appears black (Figure 1B). The chloroplasts were labeled either as having no stromules (green asterisk) or having at least one stromule (magenta asterisk). Of the 87 epidermal chloroplasts visualized, 33 have stromules. Thus, the stromule frequency in this leaf is 37.9%.
Using this analysis, the protocol was repeated for another 21 plants (one leaf per plant) during the day and a total of 24 plants at night. In the day, the average stromule frequency was 20.8 &#177; 1.8%; at night, the average stromule frequency was 12.8 &#177; 0.9% (Figure 2). Stromule frequency was significantly higher during the day, as determined by the Welch&#39;s t test (n &#8805; 22, p &#60; 0.0005). Note that although over 23,000 chloroplasts and several hundred cells were observed, the sample size, n, is reported as 22, the number of independent plants examined.
Using the protocol described here, chloroplast stromules were observed in vitro after isolation from leaves of N. benthamiana expressing plastid-targeted GFP (Figure 3A), or after using CFDA to stain chloroplasts isolated from N. benthamiana (Figure 3B) or Spinacia oleracea (Figure 3C).
<img alt="Figure 1" src="/files/ftp_upload/54692/54692fig1.jpg" /> 
Figure 1. Quantifying stromule frequency in N. benthamiana leaves. (A) A z-stack from a partial field of view of the epidermis of N. benthamiana expressing stromal GFP was merged to show all epidermal chloroplasts and stromules in a single image. (B) This image was converted to black and white and inverted for visual purposes. Chloroplasts with stromules are indicated by a magenta asterisk; chloroplasts without stromules are indicated by a green asterisk. Scale bars represent 10 &#181;m. Modified from Brunkard et al.9 <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54692/54692fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54692/54692fig2.jpg" /> 
Figure 2. Comparing stromule frequency across conditions. This protocol was used to determine the stromule frequency in 22 plants in the day and 24 plants at night. Stromule frequency was significantly higher during the day than at night (Welch&#39;s t test, n &#8805; 22, p &#60; 0.0005). Error bars indicate standard error. Modified from Brunkard et al.9 <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/54692/54692fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54692/54692fig3.jpg" /> 
Figure 3. Visualizing stromules after extracting intact chloroplasts from leaves. (A) Chloroplasts from transgenic N. benthamiana leaves expressed GFP (green) targeted to the chloroplast were isolated using the protocols described here. (B) Chloroplasts were isolated from wild-type N. benthamiana leaves and stained with CFDA, which fluoresces only inside intact, viable chloroplasts (shown in yellow). (C) Chloroplasts can also be isolated from other plant species, such as spinach (S. oleracea), and then stained with CFDA (yellow). Chlorophyll autofluorescence is shown in red. Modified from Brunkard et al.9

Watch this Scientific Journal Video about Visualizing Stromule Frequency with Fluorescence Microscopy at JoVE.com

Visualizing Stromule Frequency with Fluorescence Microscopy

Protocols to investigate the dynamics of chloroplast stromules, the stroma-filled tubules that extend from the surface of chloroplasts, are described.

可视化Stromule频率与荧光显微镜

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but ...

visualizing-stromule-frequency-with-fluorescence-microscopy

Universidad San Sebastian

Research

JoVE Journal

Biology

8.0K 次观看.  USDA. Protocols to investigate the dynamics of chloroplast stromules, the stroma-filled tubules that extend from the surface of chloroplasts, are described.

视频: Visualizing Stromule Frequency with Fluorescence Microscopy

Monitoring Spatial Segregation in Surface Colonizing Microbial Populations

Microbes provide an intriguing system to study social interaction among individuals within a population. The short generation times and relatively simple genetic modification procedures of microbes facilitate the development of the sociomicrobiology field. To assess the fitness of certain microbial species, selected strains or their genetically modified derivatives within one population, can be fluorescently labelled and tracked using microscopy adapted with appropriate fluorescence filters. Expanding colonies of diverse microbial species on agar media can be used to monitor the spatial distribution of cells producing distinctive fluorescent proteins.
Here, we present a detailed protocol for the use of green- and red-fluorescent protein producing bacterial strains to follow spatial arrangement during surface colonization, including flagellum-driven community movement (swarming), exopolysaccharide- and hydrophobin-dependent growth mediated spreading (sliding), and complex colony biofilm formation. Non-domesticated isolates of the Gram-positive bacterium, Bacillus subtilis can be utilized to scrutinize certain surface spreading traits and their effect on two-dimensional distribution on the agar-solidified medium. By altering the number of cells used to initiate colony biofilms, the assortment levels can be varied on a continuous scale. Time-lapse fluorescent microscopy can be used to witness the interaction between different phenotypes and genotypes at a certain assortment level and to determine the relative success of either.

The role of assortment (spatial segregation) in evolutionary scenarios can be examined using simple microbial systems in the laboratory that allow controlled adjustment of spatial distribution. By modifying the founder cell density, various assortment levels can be visualized using fluorescently labelled bacterial strains in the colony biofilms of Bacillus subtilis.

在表面定植微生物种群监测空间分异

Microbes provide an intriguing system to study social interaction among individuals within a population. The short generation times and relatively simple ...

科研

遗传学

Fluorescence Lifetime Imaging of Molecular Rotors in Living Cells

Diffusion is often an important rate-determining step in chemical reactions or biological processes and plays a role in a wide range of intracellular events. Viscosity is one of the key parameters affecting the diffusion of molecules and proteins, and changes in viscosity have been linked to disease and malfunction at the cellular level.1-3 While methods to measure the bulk viscosity are well developed, imaging microviscosity remains a challenge. Viscosity maps of microscopic objects, such as single cells, have until recently been hard to obtain. Mapping viscosity with fluorescence techniques is advantageous because, similar to other optical techniques, it is minimally invasive, non-destructive and can be applied to living cells and tissues.
Fluorescent molecular rotors exhibit fluorescence lifetimes and quantum yields which are a function of the viscosity of their microenvironment.4,5 Intramolecular twisting or rotation leads to non-radiative decay from the excited state back to the ground state. A viscous environment slows this rotation or twisting, restricting access to this non-radiative decay pathway. This leads to an increase in the fluorescence quantum yield and the fluorescence lifetime. Fluorescence Lifetime Imaging (FLIM) of modified hydrophobic BODIPY dyes that act as fluorescent molecular rotors show that the fluorescence lifetime of these probes is a function of the microviscosity of their environment.6-8 A logarithmic plot of the fluorescence lifetime versus the solvent viscosity yields a straight line that obeys the F&ouml;rster Hoffman equation.9 This plot also serves as a calibration graph to convert fluorescence lifetime into viscosity.
Following incubation of living cells with the modified BODIPY fluorescent molecular rotor, a punctate dye distribution is observed in the fluorescence images. The viscosity value obtained in the puncta in live cells is around 100 times higher than that of water and of cellular cytoplasm.6,7 Time-resolved fluorescence anisotropy measurements yield rotational correlation times in agreement with these large microviscosity values. Mapping the fluorescence lifetime is independent of the fluorescence intensity, and thus allows the separation of probe concentration and viscosity effects. 
In summary, we have developed a practical and versatile approach to map the microviscosity in cells based on FLIM of fluorescent molecular rotors.

Fluorescence Lifetime Imaging (FLIM) has emerged as a key technique to image the environment and interaction of specific proteins and dyes in living cells. FLIM of fluorescent molecular rotors allows mapping of viscosity in living cells.

在活细胞中的荧光寿命成像分子转子

Diffusion is often an important rate-determining step in chemical reactions or biological processes and plays a role in a wide range of intracellular ...

生物工程

Live Cell Imaging of Bacillus subtilis and Streptococcus pneumoniae using Automated Time-lapse Microscopy

During the last few years scientists became increasingly aware that average data obtained from microbial population based experiments are not representative of the behavior, status or phenotype of single cells. Due to this new insight the number of single cell studies rises continuously (for recent reviews see 1,2,3). However, many of the single cell techniques applied do not allow monitoring the development and behavior of one specific single cell in time (e.g. flow cytometry or standard microscopy).
Here, we provide a detailed description of a microscopy method used in several recent studies 4, 5, 6, 7, which allows following and recording (fluorescence of) individual bacterial cells of Bacillus subtilis and Streptococcus pneumoniae through growth and division for many generations. The resulting movies can be used to construct phylogenetic lineage trees by tracing back the history of a single cell within a population that originated from one common ancestor. This time-lapse fluorescence microscopy method cannot only be used to investigate growth, division and differentiation of individual cells, but also to analyze the effect of cell history and ancestry on specific cellular behavior. Furthermore, time-lapse microscopy is ideally suited to examine gene expression dynamics and protein localization during the bacterial cell cycle. The method explains how to prepare the bacterial cells and construct the microscope slide to enable the outgrowth of single cells into a microcolony. In short, single cells are spotted on a semi-solid surface consisting of growth medium supplemented with agarose on which they grow and divide under a fluorescence microscope within a temperature controlled environmental chamber. Images are captured at specific intervals and are later analyzed using the open source software ImageJ.

Live Cell Imaging of Bacillus subtilis and Streptococcus pneumoniae using Automated Time-lapse Microscopy

This protocol provides a step-by-step procedure to monitor single cell behavior of different bacteria in time using automated fluorescence time-lapse microscopy. Furthermore, we provide guidelines how to analyze the microscopy images.

活细胞成像枯草芽孢杆菌肺炎链球菌使用自动定时显微镜

During the last few years scientists became increasingly aware that average data obtained from microbial population based experiments are not ...

免疫与感染

High-Throughput Total Internal Reflection Fluorescence and Direct Stochastic Optical Reconstruction Microscopy Using a Photonic Chip

Total internal reflection fluorescence (TIRF) is commonly used in single molecule localization based super-resolution microscopy as it gives enhanced contrast due to optical sectioning. The conventional approach is to use high numerical aperture microscope TIRF objectives for both excitation and collection, severely limiting the field of view and throughput. We present a novel approach to generating TIRF excitation for imaging with optical waveguides, called chip-based nanoscopy. The aim of this protocol is to demonstrate how chip-based imaging is performed in an already built setup. The main advantage of chip-based nanoscopy is that the excitation and collection pathways are decoupled. Imaging can then be done with a low magnification lens, resulting in large field of view TIRF images, at the price of a small reduction in resolution. Liver sinusoidal endothelial cells (LSECs) were imaged using direct stochastic optical reconstruction microscopy (dSTORM), showing a resolution comparable to traditional super-resolution microscopes. In addition, we demonstrate the high-throughput capabilities by imaging a 500 &#181;m x 500 &#181;m region with a low magnification lens, providing a resolution of 76 nm. Through its compact character, chip-based imaging can be retrofitted into most common microscopes and can be combined with other on-chip optical techniques, such as on-chip sensing, spectroscopy, optical trapping, etc. The technique is thus ideally suited for high throughput 2D super-resolution imaging, but also offers great opportunities for multi-modal analysis.

Chip-based super-resolution optical microscopy is a novel approach to fluorescence microscopy and offers advantages in cost effectiveness and throughput. Here, the protocols for chip preparation and imaging are shown for TIRF microscopy and localization-based super-resolution microscopy.

使用光子芯片的高通量总内反射荧光和直接随机光学重建显微镜

Total internal reflection fluorescence (TIRF) is commonly used in single molecule localization based super-resolution microscopy as it gives enhanced ...

生物化学

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular level. To date the shear complexity of biological systems has caused precise single-molecule experimentation to be far too demanding, instead focusing on studies of single systems using relatively crude bulk ensemble-average measurements. However, many important processes occur in the living cell at the level of just one or a few molecules; ensemble measurements generally mask the stochastic and heterogeneous nature of these events. Here, using advanced optical microscopy and analytical image analysis tools we demonstrate how to monitor proteins within a single living bacterial cell to a precision of single molecules and how we can observe dynamics within molecular complexes in functioning biological machines. The techniques are directly relevant physiologically. They are minimally-perturbative and non-invasive to the biological sample under study and are fully attuned for investigations in living material, features not readily available to other single-molecule approaches of biophysics. In addition, the biological specimens studied all produce fluorescently-tagged protein at levels which are almost identical to the unmodified cell strains ("genomic encoding"), as opposed to the more common but less ideal approach for generating significantly more protein than would occur naturally ('plasmid expression'). Thus, the actual biological samples which will be investigated are significantly closer to the natural organisms, and therefore the observations more relevant to real physiological processes.

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Here we demonstrate the protocols for performing single-molecule fluorescence microscopy on living bacterial cells to enable functional molecular complexes to be detected, tracked and quantified.

可视化单分子络合物在体内使用高级荧光显微

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular ...

生物学

Fluorescence Time-lapse Imaging of the Complete S. venezuelae Life Cycle Using a Microfluidic Device

Live-cell imaging of biological processes at the single cell level has been instrumental to our current understanding of the subcellular organization of bacterial cells. However, the application of time-lapse microscopy to study the cell biological processes underpinning development in the sporulating filamentous bacteria Streptomyces has been hampered by technical difficulties. 
Here we present a protocol to overcome these limitations by growing the new model species, Streptomyces venezuelae, in a commercially available microfluidic device which is connected to an inverted fluorescence widefield microscope. Unlike the classical model species, Streptomyces coelicolor, S. venezuelae sporulates in liquid, allowing the application of microfluidic growth chambers to cultivate and microscopically monitor the cellular development and differentiation of S. venezuelae over long time periods. In addition to monitoring morphological changes, the spatio-temporal distribution of fluorescently labeled target proteins can also be visualized by time-lapse microscopy. Moreover, the microfluidic platform offers the experimental flexibility to exchange the culture medium, which is used in the detailed protocol to stimulate sporulation of S. venezuelae in the microfluidic chamber. Images of the entire S. venezuelae life cycle are acquired at specific intervals and processed in the open-source software Fiji to produce movies of the recorded time-series.

Fluorescence Time-lapse Imaging of the Complete S. venezuelae Life Cycle Using a Microfluidic Device

Streptomyces are characterized by a complex life cycle that has been experimentally challenging to study by cell biological means. Here we present a protocol to perform fluorescence time-lapse microscopy of the complete life cycle by growing Streptomyces venezuelae in a microfluidic device.

完整的荧光成像定时<em&gt;秒。 venezuelae</em&gt;生命周期使用微流体设备

Live-cell imaging of biological processes at the single cell level has been instrumental to our current understanding of the subcellular organization of ...

Microfluidic Tools for Probing Fungal-Microbial Interactions at the Cellular Level

Filamentous fungi are successful inhabitants of soil and play a major role in soil ecosystems, such as in the decomposition of organic and inorganic matter, as well as regulation of nutrient levels. There they also find numerous opportunities to interact with a variety of other microbes such as bacteria or other fungi. Studying fungal interactions at the cellular level, however, can be challenging owing to the black box-like nature of soil. New microfluidic tools are being developed for the study of fungal interactions; two platforms designed to study bacterial-fungal and fungal-fungal interactions are highlighted. Within these microchannels, fungal-microbial interactions can be monitored in controlled physico-chemical environments at higher temporal and spatial resolution than previously possible. Application of these tools have yielded numerous novel biological insights, such as the observation of bacterial polar attachment to hyphae or revealing uncharacterised fungal-fungal antagonisms. A key feature of these methodologies regards the ease of use of this tool by non-experts, yielding highly translatable technologies for use in microbiology labs.

Owing to the opacity of soil, interactions between its constituent microbes cannot easily be visualised with cellular resolution. Here, two microfluidic tools are presented, which offer new opportunities for investigating fungal-microbial interactions. The devices are versatile and simple to use, enabling high spatiotemporal control and high-resolution imaging at the cellular level.

用于在细胞水平上探测真菌 - 微生物相互作用的微流体工具

Filamentous fungi are successful inhabitants of soil and play a major role in soil ecosystems, such as in the decomposition of organic and inorganic ...

Temporal Quantification of MAPK Induced Expression in Single Yeast Cells

The quantification of gene expression at the single cell level uncovers novel regulatory mechanisms obscured in measurements performed at the population level. Two methods based on microscopy and flow cytometry are presented to demonstrate how such data can be acquired. The expression of a fluorescent reporter induced upon activation of the high osmolarity glycerol MAPK pathway in yeast is used as an example. The specific advantages of each method are highlighted. Flow cytometry measures a large number of cells (10,000) and provides a direct measure of the dynamics of protein expression independent of the slow maturation kinetics of the fluorescent protein. Imaging of living cells by microscopy is by contrast limited to the measurement of the matured form of the reporter in fewer cells. However, the data sets generated by this technique can be extremely rich thanks to the combinations of multiple reporters and to the spatial and temporal information obtained from individual cells. The combination of these two measurement methods can deliver new insights on the regulation of protein expression by signaling pathways.

Two complementary methods based on flow cytometry and microscopy are presented which enable the quantification, at the single cell level, of the dynamics of gene expression induced by the activation of a MAPK pathway in yeast.

在单酵母细胞MAPK诱导表达的时空定量

The quantification of gene expression at the  single cell level uncovers novel regulatory mechanisms obscured in measurements  performed at the population ...

Method for Labeling Transcripts in Individual Escherichia coli Cells for Single-molecule Fluorescence In Situ Hybridization Experiments

A method is described for labeling individual messenger RNA (mRNA) transcripts in fixed bacteria for use in single-molecule fluorescence in situ hybridization (smFISH) experiments in E. coli. smFISH allows the measurement of cell-to-cell variability in mRNA copy number of genes of interest, as well as the subcellular location of the transcripts. The main steps involved are fixation of the bacterial cell culture, permeabilization of cell membranes, and hybridization of the target transcripts with sets of commercially available short fluorescently-labeled oligonucleotide probes. smFISH can allow the imaging of the transcripts of multiple genes in the same cell, with limitations imposed by the spectral overlap between different fluorescent markers. Following completion of the protocol illustrated below, cells can be readily imaged using a microscope coupled with a camera suitable for low-intensity fluorescence. These images, together with cell contours obtained from segmentation of phase contrast frames, or from cell membrane staining, allow the calculation of the mRNA copy number distribution of a sample of cells using open-source or custom-written software. The labeling method described here can also be applied to image transcripts with stochastic optical reconstruction microscopy (STORM).

Method for Labeling Transcripts in Individual Escherichia coli Cells for Single-molecule Fluorescence In Situ Hybridization Experiments

This manuscript describes a method for labeling individual messenger RNA (mRNA) transcripts with fluorescently-labeled DNA probes, for use in single-molecule fluorescence in situ hybridization (smFISH) experiments in E. coli. smFISH is a visualization method that allows the simultaneous detection, localization, and quantification of single mRNA molecules in fixed individual cells.

单分子荧光原位杂交实验中单个大肠埃希菌细胞的转录标记方法

A method is described for labeling individual messenger RNA (mRNA) transcripts in fixed bacteria for use in single-molecule fluorescence in situ ...

Imaging Intermediate Filaments and Microtubules with 2-dimensional Direct Stochastic Optical Reconstruction Microscopy

The cytoskeleton, composed of actin microfilaments, microtubules, and intermediate filaments (IF), plays a key role in the control of cell shape, polarity, and motility. The organization of the actin and microtubule networks has been extensively studied but that of IFs is not yet fully characterized. IFs have an average diameter of 10 nm and form a network extending throughout the cell cytoplasm. They are physically associated with actin and microtubules through molecular motors and cytoskeletal linkers. This tight association is at the heart of the regulatory mechanisms that ensure the coordinated regulation of the three cytoskeletal networks required for most cell functions. It is therefore crucial to visualize IFs alone and also together with each of the other cytoskeletal networks. However, IF networks are extremely dense in most cell types, especially in glial cells, which makes its resolution very difficult to achieve with standard fluorescence microscopy (lateral resolution of ~250 nm). Direct STochastic Optical Reconstruction Microscopy (dSTORM) is a technique allowing a gain in lateral resolution of one order of magnitude. Here, we show that lateral dSTORM resolution is sufficient to resolve the dense organization of the IF networks and, in particular, of IF bundles surrounding microtubules. Such tight association is likely to participate in the coordinated regulation of these two networks and may, explain how vimentin IFs template and stabilize microtubule organization as well as could influence microtubule dependent vesicular trafficking. More generally, we show how the observation of two cytoskeletal components with dual-color dSTORM technique brings new insight into their mutual interaction.

The overall goal of this methodology is to give the optimal experimental conditions from sample preparation to image acquisition and reconstruction in order to perform 2D dual color dSTORM images of microtubules and intermediate filaments in fixed cells

可视化Stromule频率与荧光显微镜

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